1. Field of the Invention
The invention pertains to the field of X-ray crystallography. More particularly, the invention pertains to apparatuses and methods for growing crystals for X-ray crystallography.
2. Description of Related Art
Small molecule and macromolecular X-ray crystallography are a central component of modern structural genomics and drug discovery efforts. Despite the introduction of high-throughput methods, obtaining crystals of suitable size and quality for X-ray diffraction studies remains an important bottleneck in determining structures of biological macromolecules. Solution conditions (pH, salt type and concentration, protein concentration) that yield crystal growth must be identified, and then optimized to yield crystals with adequate diffraction resolution for structure determination.
Vapor diffusion is the most common method for growing crystals of proteins, viruses and biomolecular assemblies, as well as of small molecule compounds that may be useful as drugs. As shown in FIG. 8, in the manual (low-throughput) version of this technique, a drop of protein solution (50) is placed on a glass or plastic coverslip (51) that is held in an injection molded plastic frame (54). This coverslip (51) is then suspended over a reservoir (52) containing a solution (53) with a lower initial vapor pressure than that of the drop (achieved, e.g., by having a higher salt concentration or by adding polyethylene glycols.) The reservoir is sealed using clear plastic tape or a clear plastic sheet (55), and water is removed from the drop as it equilibrates with the reservoir solution, increasing the protein concentration until nucleation and crystal growth occur.
In the sitting drop method (FIG. 8(a)), the drop (50) sits on top of the supporting substrate (e.g., a coverslip (51)) and so crystals tend to sediment onto it. In the hanging drop method (FIG. 8(b)), the substrate (51) is inverted so that the drop (50) hangs from its bottom, and crystals sediment to the liquid-air interface. These hanging drops often yield crystals with unperturbed facets, show less cracking and have smaller mosaicities, and (in the absence of protein “skins”) are easily retrieved for diffraction studies. Consequently, hanging drops are preferred for final crystallization trials to obtain the largest, highest-quality crystals for data collection. In both hanging and sitting drop methods, crystals often adhere to the supporting substrate. Because protein crystals are so fragile, removing them without damaging them is often difficult.
Crystals are also grown by the batch method, where the drop is maintained at approximately uniform vapor pressure either by eliminating the reservoir solution, by choosing a reservoir solution vapor pressure equal to that of the drop, or by coating the drop in a water-impermeable oil. Since there is no water removal and associated concentration variations in time, the batch method samples only a single point in crystallization phase space. It is thus less efficient in identifying crystallization conditions than screening methods based on vapor (or liquid) diffusion.
Because of the difficulties associated with manipulating glass or plastic coverslips, especially in a high-throughput environment, injection molded multiple-cell clear plastic crystallization plates (34) (FIGS. 5(a) and 5(b)) are now widely used, especially in the initial search for solution conditions that yield crystallization. Each cell (35) in the plate (34) generally has one or more small wells (36) for the protein solution (39) and a larger well (37) for the “reservoir” or protein-free solution (38) to be equilibrated against. Solutions are dispensed into the corresponding wells, and the top of the plate is sealed using transparent tape or plastic sheets. Some manufacturers of plates for protein crystallization include Greiner Bio-One in Germany, Corning (Corning, N.Y.), Art Robbins Instruments (Sunnyvale, Calif.), Hampton Research (Aliso Viejo, Calif.) and Neuroprobe (Gaithersburg, Md.). Common drop volumes used in high throughput experiments are 2 μl or smaller. Manual crystallization to obtain large crystals for final diffraction studies may use ˜20 μl volumes.
A somewhat different crystallization plate design has been manufactured by Nextal Biotechnologies (Montreal, Canada), recently acquired by Qiagen (Venlo, Netherlands). This design is shown in FIGS. 11(a) through 11(c), and described in U.S. Patent Publication 2004/0187958. In this design, the reservoirs in the plate (70) are sealed using a screw-in cap (71) that has multiple circular ridges producing cylindrical wells (72) on its bottom (inner) side. Drops are dispensed into these cylindrical wells, and then the cap is inverted and screwed into the plate. This approach implements the hanging drop method. Light piping effects by the cylindrical walls make visualization of crystals difficult. In addition, the plate is not X-ray transparent and the walls and bottom of the cylinder provide a large surface area of plastic to which crystals may adhere, making retrieval difficult.
Protein Wave Corporation (Kyoto, Japan) has marketed thin lithographically etched sheets of an X-ray transparent polymer for use with standard crystallization plates, some aspects of which are described in U.S. Patent Publication 2003/0159641. As shown in FIGS. 12(a) and 12(b), these sheets (73) are comprised of an array of cells (74). In each cell, there is a structure with circular holes or rings (75), as well as tabs (76) that hold the structure to the sheet. These tabs allow the hole-containing structure to be easily detached from the sheet and inserted into a holder.
In use, the sheets (73) are inserted into standard multiple well plates (77) that contain the reservoir solution. Protein drops are dispensed into the circular holes (75) (instead of wells that are an integral part of the plate), and then the plate is sealed with transparent tape and equilibration of the drops and reservoir occurs. Cells that have crystals are cut open, the drop-holding structure is removed by breaking the tabs, and the structure with drops is then inserted into an X-ray beam to examine any crystals.
Because these sheets are used with standard plates, they do not allow in-situ X-ray examination. Since drops typically contain multiple crystals and large amounts of solvent, only crystal diffraction quality can be evaluated. For full molecular structure determination, crystals must still be retrieved from the drops using other tools and examined by X-rays individually. Because the drops are only supported at their edges, accelerations that occur in dispensing and routine handling lead to large drop motions and limit the volumes of liquid that can be stably supported. Consequently, a separate well is required to contain the reservoir solution with which the drops equilibrate. Because the circular rings are quite wide and are not isolated (being directly connected to the tabs and other parts of the sheet structure within the cell), the drop's contact line may occupy many positions between the inner and outer edges of the rings and even displace to other parts of the sheet structure (depending on the drop's history, its chemical composition, and its tendency to wet the sheet material), so the drop shape and position are not reproducible.
Each drop has two (top and bottom) curved, optically distorting surfaces which makes optical recognition of crystals inside difficult. Since protein “skins” (comprised of degraded or aggregated protein) often form at drop-air interfaces, the presence of two skins for these edge-supported drops makes retrieval of individual crystals for full structure analysis difficult. The instability of the edge-supported drops also makes retrieval of individual crystals for full structure determination more difficult.
Other approaches to high-throughput protein crystallization are being pursued. For example, Fluidigm (San Francisco, Calif.) has commercialized a platform based on microfluidic chips. Although these allow crystallization with very small volumes, retrieving crystals for X-ray diffraction studies is difficult, and the chips are presently very expensive compared with crystallization plates. Fluidigm's products are based on a completely different technology than the automated liquid handlers/drop dispensers that are now widely available in both academic and industrial laboratories, and do not appear to have sufficient advantages in general purpose crystallization to justify their cost.
High-throughput growth by dispensing large numbers of drops into glass or plastic capillaries is also being investigated (see, for example, B. Zheng, C. J. Gerdts and R. F. Ismagilov, Cur. Op. Struct. Biol. 15, 548-555 (2005), incorporated herein by reference), but seems unlikely to be broadly competitive with crystallization plate based methods.
Aside from these commercial technologies, there is a large body of prior art from the scientific community. Drops have been dispensed onto thin, X-ray permeable nylon loops or onto thin films and the crystals that have grown have been examined by directing X-rays through the film or loop without removing the crystals. Crystals have frequently been grown by dispensing protein and reservoir drops into X-ray transparent glass capillaries, which provide cylindrical drop support, and allow crystals to be examined by X-rays in situ. Crystals have been grown on the microfabricated grids with arrays of holes that are used in electron microscopy, including those that are X-ray permeable.
Problems with Current Technology
One of the most striking aspects of the high-throughput crystallization plates currently marketed for use with automated drop dispensing systems is how similar they all are in their basic design and function. They all have several shortcomings:
1. Most of the prior art plates implement the sitting drop technique, which yields inferior crystals to the hanging drop method. Inverting the plates to obtain hanging drops causes the contents of the reservoir and protein wells to spill out and mix.
2. Removing crystals from the plates is a difficult, time-consuming operation and at present must be done manually. Crystals often stick to the plates, and can be damaged when they are dislodged, reducing yields.
3. Although the contents of each cell can be examined optically, the protein-containing drops have irregular shapes and irreproducible positions because of their interaction with the plate surfaces. In particular, protein drops tend to be drawn to corners at the edge of the cell where they wet both the bottom and the side. This complicates automated recognition of crystals. Curved bottom wells produce more regular drops but create their own optical distortion. Flat bottom wells provide better optics but more irregularly shaped and positioned drops, and retrieving crystals from them can be more difficult.
4. The reservoir solutions are also drawn to the corners and edges of the cell, so that the shape and surface area of this solution is irregular from well to well for equal solution volumes.
5. Irregular shapes for both the protein and reservoir solutions lead to irregular surface-to-volume ratios and thus irreproducible crystallization kinetics, so that repeating and/or scaling up conditions that produced a “hit” to larger volumes to obtain larger crystals often yields no crystals.
6. Because all commercial plates are made from thick, strongly X-ray absorbing and scattering plastic, X-ray diffraction cannot be used to assess crystal quality in situ. X-rays provide the most direct method for assessing both the presence of crystals in a drop and the crystalline order of crystals present. Much time is wasted manually retrieving, mounting, flash cooling and measuring the X-ray diffraction properties of well-faceted crystals that do not diffract adequately. Although the Protein Wave design (FIG. 12) allows crystals to be grown and then extracted on an X-ray transparent holder, the process of extraction and mounting in the X-ray beam is more cumbersome than conventional methods using standard plates and separate X-ray transparent tools.
These shortcomings have a large negative impact on screening efficiency and on the overall throughput of structural genomics efforts. The average cost of determining a new protein structure is $50,000-$100,000. Given the low cost of the plates themselves compared with the costs incurred because of their deficiencies, it makes strong economic sense to replace them with a technology that is better designed to meet the demands of the high-throughput environment.